Materials and Methods
<ol>
<li>Yeast strains 
<ul>
<li>All strains used in this study are based on the BY4742 background. The SILAC control strain (Mat a, his3, leu2, ura3, lys2 and arg4::G418) was made by mating arg4 deletion strain (Mat a, his3, ura3, leu2, and arg4::G418) to BY4742 (Mat alpha, his3, leu2, ura3 and lys2), and the meiotic haploids were obtained by tetrad dissection. Therefore, the control strain is an auxotroph for lysine and arginine. The bait strain was made by PCR-mediated homologous recombination using the vector pBS1479. Therefore, the epitope tag used in this study is the tandem affinity purification (TAP) tag (3).</li>
</ul>
</li>
<li>SILAC: 
<ul>
<li>The SILAC control strain was grown in minimal dropout media supplemented with 98% L-lysine-4,4,5,5-D4 (0.03 g/L, Cambridge Isotope) and 98% L-arginine-13C6 (0.036 g/L, Cambridge Isotope).</li>
<li>The bait strain was grown in regular rich medium with normal isotopic amino acids.</li>
</ul>
</li>
<li>First stage of TAP purification [This protocol is adapted from the Cold Spring Harbor laboratory course manual (4)] **The night before your purification, pre-chill mortar and pestle in -80&deg;C freezer. Also, pre-chill a beaker in -20&deg;C** Make these buffers right before use (for 1 L of culture): 
<ul>
<li>30 ml NP-40 buffer with 25 &micro;l PIC (1/2000), 50 &micro;l of 1M DTT</li>
<li>20 ml of NP-40 buffer with 1/200 PIC, 50 &micro;l of 1M DTT</li>
<li>10 ml of TEV-C buffer with 1/2000 PIC, 10 &micro;l of 1 M DTT</li>
</ul>
</li>
</ol>
SILAC and preparation
<ol>
<li>Grow 1L each of the bait strain and the control strain separately to OD600=1.0-1.3.</li>
<li>Pour culture in 500ml Nalgene centrifuge tubes. Balance the load and pellet cells at 2580X g.</li>
<li>Decant media and resuspend pellet with 50ml of cold dH2O while transferring the cells to 50ml centrifuge tubes. Pellet cells. **You can freeze the pellet at -80&deg;C**</li>
<li>**important** match the control strain culture to the bait strain culture in 1:1 by dry pellet weight.</li>
<li>Resuspend each cell pellet in 10ml of NP-40 buffer with 1//2,000 Protease Inhibitor Cocktail (PIC).</li>
<li>Mix the two cell cultures by directly decanting one tube to the other.
 Grinding 
</li>
<li>Pour liquid N2 into the pre-chilled mortar and allow it to completely evaporate. Add 2 ml of cells to the mortar in circular motion. Pour some liquid N2 to freeze the cells. Grind the cells until the cells become fine powder. Repeat the process until all the cells are ground. **Do not allow the cells to thaw. Add liquid N2 when necessary**</li>
<li>Transfer the powder to an ice-cold beaker and thaw at room temperature. When the edges begin to thaw, add 20 ml of NP-40 buffer with 1/200 PIC.</li>
<li>Transfer crude lysate to 40-ml Nalgene tubes. Spin at 39000 xgfor 30 min.</li>
<li>While waiting, take 300 &mu;l of Sepharose 2B beads (GE). Wash the beads in 300 &mu;l of NP-40 buffer (1/2,000 PIC) 3 times. Resuspend the beads in 300 &mu;l buffer so that they are in 1:1 slurry.
 Pre-clearing cell lysate 
</li>
<li>Transfer the supernatant to a new tube and add the Sepharose 2B beads. Incubate on a rotating platform in cold room for 30 min.</li>
<li>While waiting, take 300 &mu;l of IgG Sepharose 6 beads (GE). Wash the beads in 300 &mu;l of NP-40 buffer (1/2,000 PIC) 3 times. Resuspend the beads in 300 _l buffer so that they are in 1:1 slurry.</li>
<li>Pellet the beads. Transfer the supernatant to a new tube. **Save an aliquot of the cell lysate for Western Blotting later**
 Binding to IgG sepharose beads 
</li>
<li>Remove the Sepharose 2B beads by centrifuge. Add the IgG beads (previously prepared in 1:1 slurry). Separate the contents into two tubes for more efficient binding. Incubate on a rotating platform in cold room for 2 hr.</li>
<li>Spin down the beads. **Save an aliquot of the unbound faction**</li>
<li>Wash beads with 10 ml NP-40 buffer (1/2,000 PIC).</li>
<li>Wash beads with 3 ml TEV-C buffer (1/2,000 PIC). **Save an aliquot of the beads. This will reflect the efficiency of binding**
 Eluting from IgG beads by TEV protease cleavage 
</li>
<li>Add 5 &mu;l of AcTEV to 1ml of TEV-C buffer (with 1/2,000 PIC). After mixing, separate the contents into two 1.5 ml Eppendorf tubes for more efficient mixing. Incubate on a rotating platform in cold room overnight.</li>
<li>Spin down the beads and transfer the eluate to a fresh 1.5-ml tube. Wash the beads with additional 0.5 ml of TEV-C buffer</li>
<li>Combine the elaute in one tube. You should have 1.5 ml of eluate in total. **Save an aliquot of the TEV eluate**
 Trichloroacetic acid (TCA) precipatation (For mass spectrometry-based analysis, we recommend using EtOH/ acetate precipitation) 
</li>
<li>Adjust aliquots to 25% TCA with 100% TCA. To do this, separate the 1.5 ml eluate into 2 tubes of 750 &mu;l each. Add 250 &mu;l of 100% TCA to the tube. Your solution should turn milky.</li>
<li>Place on ice for 30 min with periodic vortexing.</li>
<li>Spin at maximum speed in the table-top centrifuge in cold room for 30 min.</li>
<li>Wash once with 500 &mu;l of ice-cold acetone containing 0.05 N HCl and spin for 5 min at maximum speed (cold room).</li>
<li>Wash once with 500 &mu;l of ice-cold acetone and spin for 5 min at max (cold room).</li>
<li>Carefully remove acetone and dry pellet.</li>
<li>For silver staining, resuspend the pellet in 50 &mu;l of 1X SDS sample buffer.
 EtOH/ acetate precipatation 
</li>
<li>Add 20 &micro;g of glycogen</li>
<li>Dilute samples 5X with 100% EtOH and adjust to 50 mM NaCH3COO with a 2.5 M stock, pH 5.0</li>
<li>Stand the solution for 2hr</li>
<li>Pellet precipitated protein by centrifugation for 10 min at 12,000 X g at room temperature. </li>
</ol>
NP-40 Buffer (for 200 ml)
<table border="0">
<tbody>
<tr>
<td>&nbsp;</td>
<td>Stock concentration</td>
<td>Add</td>
</tr>
<tr>
<td>10 mM sodium phosphate buffer (pH 7.2)</td>
<td>0.1 M</td>
<td>20 ml</td>
</tr>
<tr>
<td>150 mM NaCl</td>
<td>2 M</td>
<td>15 ml</td>
</tr>
<tr>
<td>1% NP-40</td>
<td>100%</td>
<td>2 ml</td>
</tr>
<tr>
<td>50 mM NaF</td>
<td>1 M</td>
<td>10 ml</td>
</tr>
<tr>
<td>0.1 mM Na3VO4</td>
<td>10 mM</td>
<td>2 ml</td>
</tr>
<tr>
<td style="text-align:right;" colspan="3">Volume to 200 ml</td>
</tr>
</tbody>
</table>
ADD before use:
<ol>
<li>1/1,000 of 1 M DTT</li>
<li>1/2,000 of PIC (Protease Inhibitors Cocktail)</li>
</ol>
TEV-C Buffer (for 50 ml)
<table border="0">
<tbody>
<tr>
<td>&nbsp;</td>
<td>Stock concentration</td>
<td>Add</td>
</tr>
<tr>
<td>25 mM Tris (pH 8.0)</td>
<td>100 mM</td>
<td>12.5 ml</td>
</tr>
<tr>
<td>150 mM NaCl</td>
<td>2 M</td>
<td>3.75 ml</td>
</tr>
<tr>
<td>0.1 % NP-40</td>
<td>100 %</td>
<td>50 _l</td>
</tr>
<tr>
<td>0.5 mM EDTA</td>
<td>500 mM</td>
<td>50 __l</td>
</tr>
<tr>
<td style="text-align:right;" colspan="3">Volume to 50 ml</td>
</tr>
</tbody>
</table>
ADD before use:
<ol>
<li>1/1,000 of 1M DTT</li>
<li>1/2,000 of PIC</li>
</ol>

<ol>
	<li>Loewen, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inheritance+of+cortical+ER+in+yeast+is+required+for+normal+septin+organization.">Inheritance of cortical ER in yeast is required for normal septin organization.</a> Jour. Cell Biol. 179, 467-483 .</li><li>Ong, S., Foster, L. J., Hoog, C. L., Mann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mass+spectrometric-based+approaches+in+quantitative+proteomics.">Mass spectrometric-based approaches in quantitative proteomics.</a> Methods. 29, 124-130 .</li><li>Puig, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Tandem+Affinity+Purification+(TAP)+method:+a+general+procedure+of+protein+complex+purification.">The Tandem Affinity Purification (TAP) method: a general procedure of protein complex purification.</a> Methods. 24, 218-229 .</li><li>Amberg, D. C., Burke, D. J., Strathern, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+in+Yeast+Genetics.">Methods in Yeast Genetics.</a> , 123-125 .</li></ol>

The aliquots saved during the purification procedure should include (1) pre-cleared cell lysate, (2) bound fraction, (3) unbound fraction, and (4) eluted fraction. We recommend analyzing the protein contents of the above aliquots by Western blotting using anti-TAP antibody or silver staining to reflect the binding and eluting efficiency of the experiment. Examples of a silver stained gel and a blot are shown in Figure 1.
Since SILAC provides us with unbiased and quantified measurements of protein binding partners, we only do the first stage of the TAP purification to minimize sample loss. If you experience significant amounts of unspecific binding, you may wish to carry out the entire protocol, which can be found in the Cold Spring Harbor manual (4).

identification of protein complexes with quantitative proteomics in s. cerevisiae

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as important intracellular signalling molecules. However, unlike proteins, lipids are small hydrophobic molecules that traffic primarily by poorly described nonvesicular routes, which are hypothesized to occur at membrane contact sites (MCSs). MCSs are regions where the endoplasmic reticulum (ER) makes direct physical contact with a partnering organelle, e.g., plasma membrane (PM). The ER portion of ER-PM MCSs is enriched in lipid-synthesizing enzymes, suggesting that lipid synthesis is directed to these sites and implying that MCSs are important for lipid traffic. Yeast is an ideal model to study ER-PM MCSs because of their abundance, with over 1000 contacts per cell, and their conserved nature in all eukaryotes. Uncovering the proteins that constitute MCSs is critical to understanding how lipids traffic is accomplished in cells, and how they act as signaling molecules. We have found that an ER called Scs2p localize to ER-PM MCSs and is important for their formation. We are focused on uncovering the molecular partners of Scs2p. Identification of protein complexes traditionally relies on first resolving purified protein samples by gel electrophoresis, followed by in-gel digestion of protein bands and analysis of peptides by mass spectrometry. This often limits the study to a small subset of proteins. Also, protein complexes are exposed to denaturing or non-physiological conditions during the procedure. To circumvent these problems, we have implemented a large-scale quantitative proteomics technique to extract unbiased and quantified data. We use stable isotope labeling with amino acids in cell culture (SILAC) to incorporate staple isotope nuclei in proteins in an untagged control strain. Equal volumes of tagged culture and untagged, SILAC-labeled culture are mixed together and lysed by grinding in liquid nitrogen. We then carry out an affinity purification procedure to pull down protein complexes. Finally, we precipitate the protein sample, which is ready for analysis by high-performance liquid chromatography/ tandem mass spectrometry. Most importantly, proteins in the control strain are labeled by the heavy isotope and will produce a mass/ charge shift that can be quantified against the unlabeled proteins in the bait strain. Therefore, contaminants, or unspecific binding can be easily eliminated. By using this approach, we have identified several novel proteins that localize to ER-PM MCSs. Here we present a detailed description of our approach. 

Watch this Scientific Journal Video about Identification of protein complexes with quantitative proteomics in S. cerevisiae at JoVE.com

Identification of protein complexes with quantitative proteomics in S. cerevisiae

Here we describe a new quantitative proteomics technique for identifying protein complexes in Saccharomyces cerevisiae. In this study, we have used the SILAC method together with affinity purification followed by tandem mass spectrometry to identify with high specificity the binding partners of an ER protein, Scs2p.

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as ...

identification-protein-complexes-with-quantitative-proteomics-s

Nanyang Technological University

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13.1K Views.  University of British Columbia - UBC. Here we describe a new quantitative proteomics technique for identifying protein complexes in Saccharomyces cerevisiae. In this study, we have used the SILAC method together with affinity purification followed by tandem mass spectrometry to identify with high specificity the binding partners of an ER protein, Scs2p.

Video: Identification of protein complexes with quantitative proteomics in S. cerevisiae

Assay for Adhesion and Agar Invasion in S. cerevisiae

Yeasts are found in natural biofilms, where many microorganisms colonize surfaces. In artificial environments, such as surfaces of man-made objects, biofilms can reduce industrial productivity, destroy structures, and threaten human life. 1-3 On the other hand, harnessing the power of biofilms can help clean the environment and generate sustainable energy. 4-8
The ability of S. cerevisiae to colonize surfaces and participate in complex biofilms was mostly ignored until the rediscovery of the differentiation programs triggered by various signaling pathways and environmental cues in this organism. 9, 10 The continuing interest in using S. cerevisiae as a model organism to understand the interaction and convergence of signaling pathways, such as the Ras-PKA, Kss1 MAPK, and Hog1 osmolarity pathways, quickly placed S. cerevisiae in the junction of biofilm biology and signal transduction research. 11-20 To this end, differentiation of yeast cells into long, adhesive, pseudohyphal filaments became a convenient readout for the activation of signal transduction pathways upon various environmental changes. However, filamentation is a complex collection of phenotypes, which makes assaying for it as if it were a simple phenotype misleading. In the past decade, several assays were successfully adopted from bacterial biofilm studies to yeast research, such as MAT formation assays to measure colony spread on soft agar and crystal violet staining to quantitatively measure cell-surface adherence. 12, 21 However, there has been some confusion in assays developed to qualitatively assess the adhesive and invasive phenotypes of yeast in agar.
Here, we present a simple and reliable method for assessing the adhesive and invasive quality of yeast strains with easy-to-understand steps to isolate the adhesion assessment from invasion assessment. Our method, adopted from previous studies, 10, 16 involves growing cells in liquid media and plating on differential nutrient conditions for growth of large spots, which we then wash with water to assess adhesion and rub cells completely off the agar surface to assess invasion into the agar. We eliminate the need for streaking cells onto agar, which affects the invasion of cells into the agar. In general, we observed that haploid strains that invade agar are always adhesive, yet not all adhesive strains can invade agar medium. Our approach can be used in conjunction with other assays to carefully dissect the differentiation steps and requirements of yeast signal transduction, differentiation, quorum sensing, and biofilm formation.

We describe a qualitative assay for yeast adhesion and agar invasion as a measure of invasive and pseudohyphal differentiation. This simple assay can be used to assess the invasive phenotype of various mutants as well as the effects environmental cues and signaling pathways on yeast differentiation.

Yeasts are found in natural biofilms, where many microorganisms colonize surfaces. In artificial environments, such as surfaces of man-made objects, ...

Proteomics to Identify Proteins Interacting with P2X2 Ligand-Gated Cation Channels

Ligand-gated ion channels underlie synaptic communication in the nervous system1. In mammals there are three families of ligand-gated channels: the cys loop, the glutamate-gated and the P2X receptor channels2. In each case binding of transmitter leads to the opening of a pore through which ions flow down their electrochemical gradients. Many ligand-gated channels are also permeable to calcium ions3, 4, which have downstream signaling roles5 (e.g. gene regulation) that may exceed the duration of channel opening. Thus ligand-gated channels can signal over broad time scales ranging from a few milliseconds to days. Given these important roles it is necessary to understand how ligand-gated ion channels themselves are regulated by proteins, and how these proteins may tune signaling. Recent studies suggest that many, if not all, channels may be part of protein signaling complexes6. In this article we explain how to identify the proteins that bind to the C-terminal aspects of the P2X2 receptor cytosolic domain.
P2X receptors are ATP-gated cation channels and consist of seven subunits (P2X1-P2X7). P2X receptors are widely expressed in the brain, where they mediate excitatory synaptic transmission and presynaptic facilitation of neurotransmitter release7. P2X receptors are found in excitable and non-excitable cells and mediate key roles in neuronal signaling, inflammation and cardiovascular function8. P2X2 receptors are abundant in the nervous system9 and are the focus of this study. Each P2X subunit is thought to possess two membrane spanning segments (TM1 &amp; TM2) separated by an extracellular region7 and intracellular N and C termini (Fig 1a)7. P2X subunits10 (P2X1-P2X7) show 30-50% sequence homology at the amino acid level11. P2X receptors contain only three subunits, which is the simplest stoichiometry among ionotropic receptors. The P2X2 C-terminus consists of 120 amino acids (Fig 1b) and contains several protein docking consensus sites, supporting the hypothesis that P2X2 receptor may be part of signaling complexes. However, although several functions have been attributed to the C-terminus of P2X2 receptors9 no study has described the molecular partners that couple to the intracellular side of this protein via the full length C-terminus. In this methods paper we describe a proteomic approach to identify the proteins which interact with the full length C-terminus of P2X2 receptors.

We describe a simple protocol to identify brain proteins that bind to the full length C terminus of ATP-gated P2X2 receptors. The extension and systematic application of this approach to all P2X receptors is expected to lead to a better understanding of P2X receptor signaling.

Ligand-gated ion channels underlie synaptic communication in the nervous system1. In mammals there are three families of ligand-gated channels: the cys ...

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground using a cryolysis procedure in a ball mill grinder and the resulting grindate brought into solution by urea solubilization. This procedure allows for the lysis of the cells in a metabolically inactive state, preserving phosphorylation and preventing reorientation of the phosphoproteome during cell lysis.  Following reduction, alkylation, trypsin digestion of the proteins, the samples are desalted on C18 columns and the sample complexity reduced by fractionation using hydrophilic interaction chromatography (HILIC). HILIC columns preferentially retain hydrophilic molecules which is well suited for phosphoproteomics.  Phosphorylated peptides tend to elute later in the chromatographic profile than the non phosphorylated counterparts. After fractionation, phosphopeptides are enriched using immobilized metal chromatography, which relies on charge-based affinities for phosphopeptide enrichment. At the end of this procedure the samples are ready to be quantitatively analyzed by mass spectrometry.

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

Description of a quantitative phosphorylation procedure using cryolysis, urea solubilziation, HILIC fractionation and IMAC enrichment of phosphorylated peptides.

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

Identification of Post-translational Modifications of Plant Protein Complexes

Plants adapt quickly to changing environments due to elaborate perception and signaling systems. During pathogen attack, plants rapidly respond to infection via the recruitment and activation of immune complexes. Activation of immune complexes is associated with post-translational modifications (PTMs) of proteins, such as phosphorylation, glycosylation, or ubiquitination. Understanding how these PTMs are choreographed will lead to a better understanding of how resistance is achieved.
Here we describe a protein purification method for nucleotide-binding leucine-rich repeat (NB-LRR)-interacting proteins and the subsequent identification of their post-translational modifications (PTMs). With small modifications, the protocol can be applied for the purification of other plant protein complexes. The method is based on the expression of an epitope-tagged version of the protein of interest, which is subsequently partially purified by immunoprecipitation and subjected to mass spectrometry for identification of interacting proteins and PTMs.
This protocol demonstrates that: i). Dynamic changes in PTMs such as phosphorylation can be detected by mass spectrometry; ii). It is important to have sufficient quantities of the protein of interest, and this can compensate for the lack of purity of the immunoprecipitate; iii). In order to detect PTMs of a protein of interest, this protein has to be immunoprecipitated to get a sufficient quantity of protein.

We describe here a protocol for the purification and characterization of plant protein complexes. We demonstrate that by immunoprecipitating a single protein within a complex, so we can identify its post-translational modifications and its interacting partners.

Plants adapt quickly to changing environments due to elaborate perception and signaling systems. During pathogen attack, plants rapidly respond to ...

Identification of Protein Interaction Partners in Mammalian Cells Using SILAC-immunoprecipitation Quantitative Proteomics

Quantitative proteomics combined with immuno-affinity purification, SILAC immunoprecipitation, represent a powerful means for the discovery of novel protein:protein interactions. By allowing the accurate relative quantification of protein abundance in both control and test samples, true interactions may be easily distinguished from experimental contaminants. Low affinity interactions can be preserved through the use of less-stringent buffer conditions and remain readily identifiable. This protocol discusses the labeling of tissue culture cells with stable isotope labeled amino acids, transfection and immunoprecipitation of an affinity tagged protein of interest, followed by the preparation for submission to a mass spectrometry facility. This protocol then discusses how to analyze and interpret the data returned from the mass spectrometer in order to identify cellular partners interacting with a protein of interest. As an example this technique is applied to identify proteins binding to the eukaryotic translation initiation factors: eIF4AI and eIF4AII.

SILAC immunoprecipitation experiments represent a powerful means for discovering novel protein:protein interactions. By allowing the accurate relative quantification of protein abundance in both control and test samples, true interactions may be easily distinguished from experimental contaminants, and low affinity interactions preserved through use of less-stringent buffer conditions.

Quantitative proteomics combined with immuno-affinity purification, SILAC immunoprecipitation, represent a powerful means for the discovery of novel ...

Identification of protein complexes with quantitative proteomics in S. cerevisiae

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Identification of Post-translational Modifications of Plant Protein Complexes

Identification of Protein Interaction Partners in Mammalian Cells Using SILAC-immunoprecipitation Quantitative Proteomics